Hydroxylated Bisabolol Oxides: Evidence for Secondary Oxidative

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Hydroxylated Bisabolol Oxides: Evidence for Secondary Oxidative Metabolism in Matricaria chamomilla Cristina Avonto,† Mei Wang,† Amar G. Chittiboyina,† Bharathi Avula,† Jianping Zhao,† and Ikhlas A. Khan*,†,‡ †

National Center for Natural Products Research, The University of Mississippi, Mississippi 38677, United States Department of Pharmacognosy, School of Pharmacy, The University of Mississippi, Mississippi 38677, United States

‡

S Supporting Information *

ABSTRACT: German chamomile (Matricaria chamomilla) is one of the most popular medicinal plants used in Western herbal medicine. Among the various phytochemicals present in the essential oil of the flowers of German chamomile, bisabolol and its oxidized metabolites are considered as marker compounds for distinguishing different chemotypes. These compounds are influential in mediating the aroma of the essential oil of M. chamomilla and contribute to the therapeutic properties (anti-inflammatory, antibacterial, insecticidal, and antiulcer) of this species. In order to find other possible bisabolol derivatives as marker compounds for authentication of German chamomile in botanical and commercial products, an in-depth investigation using a GC-assisted fractionation procedure was performed on nonpolar fractions. As a result of this approach, three new hydroxylated derivatives of bisabolol oxides A and B (1−3) have been isolated from M. chamomilla. Plausible biogenetic pathways are presented.

G

bisabolol has Generally Regarded as Safe (GRAS) status,12 and it is responsible for many beneficial effects of German chamomile essential oil. α-Bisabolol has been found to have anti-inflammatory,13 antibacterial and antifungal,14 insecticidal,15 and antiulcer properties.16 Also, this compound was shown to possess a promising potential selective anticancer activity, by inducing apoptosis in malignant glioblastoma tumor cells.17 Bisabolol oxides A and B also contribute to the overall antispasmodic effect of chamomile preparations.18 As part of ongoing research on the authentication, validation, safety and biological evaluation of dietary supplements,19 German chamomile was selected for further comprehensive analysis. In order to find other possible minor volatile and nonvolatile bisabolol-type compounds as authentication markers for M. chamomilla, an in-depth investigation of minor compounds from this species has been undertaken. A gas chromatography-assisted fractionation approach was used in order to identify bisabolol derivatives occurring in low concentrations among the several classes of compounds found in the complex chamomile extract.

erman chamomile, Matricaria chamomilla L. (Asteraceae), is one of the most important medicinal plants in Western herbal medicine. It has been used as a traditional remedy for more than 2500 years and reported as a panacea by several renowned physicians such as Hippocrates, Galen, and Asclepius.1 In modern times, chamomile traditional uses have been validated by a wide number of scientific investigations of its biological activity.2 On the basis of these facts, a plethora of commercial products have been produced containing chamomile extracts, including herbal teas, hair products, soaps, lotions, creams, and perfumes. Chemically, the majority of active compounds are produced mainly in the flower-head parts of M. chamomilla.3 So far, more than 200 known compounds have been isolated and identified, including volatile terpenoids,4 flavonoids and their glycoside derivatives,5 coumarins,6 fatty acids,7 alkaloids,8 and polysaccharides.9 This chemical diversity is responsible for the wide variety of biological activities that have been attributed to both the essential oil and the aqueous extract10 of German chamomile. Six different chemotypes of M. chamomilla have been defined according to the chemical composition of the essential oil.3 More than 60 sesquiterpenoids have been identified among volatile compounds from M. chamomilla essential oils,11 including linear, guaiane-type, and bisabolane-type sesquiterpenoids. α-Bisabolol, bisabolol oxides A and B, and α-bisabolone oxide A are used as marker compounds for chemotype identification and authentication of products containing German chamomile.3 While α-bisabolol is commonly found in a wide variety of plants, its furano and pyrano derivatives (bisabolol oxides) are less common natural products, but all these compounds are of potential pharmacological importance. For example, in the United States, α© XXXX American Chemical Society and American Society of Pharmacognosy

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RESULTS AND DISCUSSION German chamomile crude extracts exhibited very complex chemical profiles, making the purification of minor metabolites a time-consuming and costly process. The hexane fraction obtained from liquid−liquid partitioning of M. chamomilla flowers contained most of the compounds of interest among a large number of fatty acids, their methyl esters, polyacetylene Received: April 23, 2013

A

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relationship between the two compounds. The configuration at the new stereocenter (C-9) was confirmed by NOESY experiments, in which the signals of H-8a and H-8b showed correlations with both the H-6 and the C-15 methyl group signal, respectively. The H-6 signal gave a correlation with H-8a while showing no correlation with H-7a and H-7b. On the basis of these observations, the relative configuration at C-9 was assigned as shown. These data led to the characterization of this new compound as 9-hydroxybisabolol oxide (1). In the 13C NMR spectrum of compound 2, the presence of 15 signals was evident, of which five were identified from the DEPT-135 spectrum as methylenes, with four methyl groups, four quaternary carbons, and two methine carbons also apparent. The chemical shifts for the two carbons at δ 144.1 and 112.4 ppm were assigned to a monosubstituted terminal alkene, which was confirmed by the splitting pattern and COSY correlation of the doublet of doublets at δ 5.90 with two doublets at δ 5.23 and 5.04 ppm, respectively. The two sp3 carbons with chemical shifts at δ 72.1 and 70.7 were assigned to two tertiary carbons attached to hydroxy groups (C-3 and C-11, respectively). Both the vinylic protons and the methyl group at C-15 gave HMBC correlations with the quaternary carbon at δ 72.1 ppm. The carbonyl 13C NMR signal at δ 211.5 (C-5) gave a HMBC correlation with the diastereotopic protons at δ 2.84− 2.72 and δ 2.67−2.59 ppm, respectively, thus placing the carbonyl in a beta position to the quaternary hydroxy group at δ 72.1. Two additional signals at δ 82.5 (C-7) and 86.4 ppm (C-10) suggested the presence of a tetrahydrofuran moiety. Additionally, gHMBC correlations of two methyl groups at δ 1.53 and 1.52 ppm were assigned to the isopropyl moiety connected at the C-10 position. Connections of the methyl group at δ 1.28 ppm with C-7 also confirmed the structural relationship of the tetrahydrofuran ring with the B-ring of bisabolol oxide B. In the positive HRESIMS, a peak occurred for a [M − OH]+ ion at m/z 253.1785 (calcd m/z 253.1799) instead of m/z 270.1831 expected for the parent compound, probably due to loss of H2O. The new compound isolated was identified as seco-bisabolol oxide B (2). For the third new compound (3), the 1H and 13C NMR chemical shifts were closely related to those of the known compound bisabolol oxide A glycoside (4),21 which was also isolated from German chamomile. All the main signals were thus assigned based on the parent compound bisabolol oxide A glucoside.20,21 By comparison with compound 4, the DEPT135 and 13C data showed the presence of an additional CH2 signal at δ 64.4 ppm, while lacking one of the methyl singlets of 4. From gHMBC and COSY correlations this signal was therefore assigned to the methylene at C-15. The D-glucose moiety was confirmed by both acid and enzymatic hydrolysis followed by chemical derivatization and comparison with standards of D- and L-glucose using GC-MS. Spectroscopic data confirmed the β linkage between the aglycone and the glucose moiety [C-1′ δ 103.50 ppm, H-1′ (d, J = 7.8 Hz)]. NOESY correlations between H-12 and both H-14 and H-2 confirmed the relative configuration of the three stereocenters of the aglycone, which retained the same configuration of compound 4. A sodiated molecular ion peak was observed in the HRESIMS at m/z 439.2296. This new compound was characterized as 15-hydroxybisabolol oxide A glycoside (3). Biogenetically, compounds 1−5 can be interconnected through the commonly accepted bisabolane biosynthetic pathway. Compounds 1, 3, and 5 could be derived from allylic hydroxylation of the parent terpenoid α-bisabolol. This step

derivatives, and other volatile sesquiterpenes not related to bisabolol metabolites. In order to find a robust method for the isolation of bisabolol-like compounds, the nonpolar mixture obtained from partitioning of the flower-head methanol extract was subjected to fractionation, and all fractions generated were analyzed by GC-MS. The mass spectrometric profile was matched with the comprehensive NIST 2008 chemical compound database. Bisabolol oxides A and B possess a very distinctive fragmentation pattern, with a base peak at m/z 143 and secondary fragmentation peaks at m/z 125, 107, and 71 (see Scheme S1, Supporting Information, for a proposed fragmentation mechanism leading to generation of the significant peaks). This fragmentation pattern provided an easy and convenient tool for detection of bisabolol oxide-like compounds in the crude extract. Further purification focused mainly on fractions containing peaks of interest that conformed to this characteristic fragment pattern, thus leading to the focused isolation of three new compounds (1−3) and one known compound (5) never previously reported from German chamomile. Compound 1 was isolated as a pale yellow oil. The observed pseudomolecular ion at m/z 272.2199 was obtained by HRESIMS (calcd for [C15H26O3 + NH4]+, 272.2221). The molecular formula suggested three degrees of unsaturation. The 1D NMR data and EIMS fragmentation profile were compatible with the presence of a hydroxypyrano moiety related to the B-ring of bisabolol oxide A. The 13C NMR spectroscopic data (DEPT-135 and 13C) showed 15 signals, which were assigned to one double bond (δ 134.9 and 131.0 ppm, respectively), four oxygenated carbons (between δ 75.1 and 67.3), one tertiary carbon at δ 48.2, and eight signals in the range δ 37.4−19.9 ppm (four methyl and four methylene moieties). The disubstituted olefin was confirmed by 1H NMR and COSY correlation of the 1H NMR signals at δ 5.89 and 5.75 ppm. The presence of four isolated methyl groups was determined by the four singlets between δ 1.28 and 1.22 ppm. Final connectivities as shown in Figure S2, Supporting Information, were assigned by gradient-HMQC, 1H−1H COSY, and gradient-HMBC NMR experiments. The positions of the hydroxy group and the double bond were established with gHMBC correlation peaks of H3-15 with the vinyl carbon at C-10, the oxygenated carbon C-9, and the methylene moiety at the C-8 position. Altogether the NMR data of 1 were similar to those of bisabolol oxide A,20 suggesting a close structural B

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Scheme 1. Proposed Enzymatic Oxidation of Bisabolol Oxide Derivatives 1, 3, and 5a

a

The allylic hydroxylation step can occur before or after ring B closure and is eventually followed by further glycosylation steps.

Scheme 2. Proposed Pathway That Leads to the Generation of 2a

a

The loss of the pyrophosphate group and migration of the double bond would result in the formation of 2a, which upon water addition would provide 2b. Allylic hydroxylation and further oxidation would lead to the formation of a keto group at the C-5 position. Protonation and addition of water to the resulting tertiary carbocation would generate the hydroxy group at the C-8 position. Epoxidation (b) and ring-opening (c, d) by intramolecular nucleophilic substitution is a generally accepted mechanism for formation of the tetrahydrofuran ring of bisabolol oxide B.

could possibly occur through enzymatic transformation because only one of two possible isomers has been identified in any of these compounds. This can occur either before or after the epoxidation and cyclization reactions that lead to bisabolol oxides A and B (Scheme 1). Such kinds of enzymatic modification are well-known in plant metabolism. Whereas the biosynthetic routes for sesquiterpenes are not fully understood, some mechanistic details are available for monoterpene syntheses, and cytochrome P450 is known to be involved in the hydroxylation of monoterpenes.22 The aglycone moiety of 3 was not found in the nonpolar fractions of M. chamomilla flowers, and this fact was attributed to either low abundance or the instability/volatility characteristics of the compound during the isolation procedure, but its presence cannot be excluded. Hydroxylated bisabolol oxides rarely occur in Nature, but some examples have been reported. For example, the 15-hydroxy derivative of α-bisabolol has been isolated from Vanillosmopsis arborea (Gardner) Baker (Compositae);23 this supports the possibility of the occurrence of the parent aglycone of 3 in German chamomile flowers. Glycosylated derivatives are naturally occurring in chamomile, as demonstrated by the isolation of bisabolol oxide A glucoside (4). Both compound 5 and the 2-acetoxy derivative of 1 are known metabolites from other members of the Compositae family, e.g., Artemisia lucentica O. Bolòs, Vallès & Vigo.24

Bisabolol and its derivatives are characterized by the presence of the A-ring attached to a five-membered or six-membered ether linkage (B-ring). Similar to bisabolol oxide B, compound 2 contains a furano ring but lacks the essential six-membered ring A. The commonly accepted pathway for bisabolane compounds involves the formation of the A-ring followed by further enzymatic steps that lead to formation of the pyrano or furano moiety.25 Compound 2 is unlikely to be an isolation artifact, suggesting that this compound may originate from a different biosynthetic pathway, wherein hydroxylation of the first prenyl unit occurs as a first step, leading to nerolidol. This is plausible, since all the bisabolane sesquiterpenoids are derived from the nerolidyl diphosphate pathway.26 Due to low abundance and compound instability, elucidation of the configuration at C-3 was not feasible in this study. Nonetheless, most of the enzymes involved in the nerolidol synthesis show good enantioselectivity toward the 3S-stereoisomer,27 thus the S-configuration at the C-3 position of 2 is most probable assuming that biosynthesis follows the nerolidol synthesis. Moreover, to support this hypothesis, a similar sesquiterpenoid named neroplofurol has been recently isolated for the first time from Oplopanax horridus (Sm.) Miq.28 The proposed biosynthetic pathway for neroplofurol involves epoxidation at the prenyl units and subsequent cascade ring-opening due to addition of water.29 In the case of compound 2, this mechanism is unlikely because of C

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final temperature of 280 °C (held for 30 min). The column used for the analysis was an Agilent HP-5MS GC capillary column (30 m × 0.25 mm × 0.25 μm). The electron-impact ion source was set to 70 eV. The mass spectrometer was scanned from 40 to 550 amu. ChemStation software was used for acquisition, processing, and calibration of the GC-MS data. The major compounds were identified by comparing the retention times with the reference standards where possible, reported literature data, and NIST library searches. Flash column chromatography purifications were performed on Biotage Isolera Four, using SNAP cartridges or Flash+ HPFC cartridges. Silica gel cartridges were KP-Sil (40−63 μm, 60 Å) or KP-C18-HS (35−70 μm, 90 Å), 18% carbon load (by weight). Gravity column chromatography (CC) purifications were performed using silica gel (40−63 μm, 60 Å, Sorbtech), reversed-phase RP C18 silica (Polarbond, J.T. Baker), neutral alumina oxide (50−200 μm, Acrös Organic), and Sephadex LH-20 (Sigma-Aldrich). β-Glucosidase from almonds (10−30 units/mg solid, CAS 9001-223, Sigma-Aldrich) was used for enzymatic hydrolysis. Disodium hydrogen phosphate-citric acid (DHPCA) buffer solution was prepared with citric acid 0.1 M and Na2HPO4 0.2 M, and the final pH adjusted to 5.2. Plant Material. M. chamomilla flower heads were supplied by Frontier Natural Products Co-op. (Norway, IA, USA). Plant material was authenticated by Dr. Vijayasankar Raman at the National Center for Natural Products Research (NCNPR), The University of Mississippi, Mississippi, United States. A sample specimen (voucher number 9365) is available at the National Center for Natural Product Research, University of Mississippi. Extraction and Isolation. Air-dried M. chamomilla flower heads (840 g) were finely powered and extracted three times with MeOH (3 L) by sonication (30 min). Solid particles were then separated by filtration, and the solvent was evaporated under vacuum at temperatures below 40 °C. The crude extract thus obtained (171.0 g) was then suspended in water (500 mL) and sequentially partitioned with hexane, chloroform, ethyl acetate, and n-butanol. The fractions obtained weighed 31.1, 9.4, 11.6, and 39.8 g, respectively. The hexane fraction was separated on silica gel, and the subfractions collected were combined into 20 main fractions (A1−A20). Isolation of unknown compounds from the hexane fraction was performed using a GC-assisted approach. The crude hexane fraction and all obtained fractions were analyzed by GC-MS. Compound identification was achieved by comparing the spectra with the NIST database using a probability-based matching algorithm. Compounds with a similar structure to bisabolol oxides A and B exhibited a similar fragmentation pattern, with a base peak at m/z 143 and additional fragmentation peaks at m/z 125, 107, and 71. This pattern was considered suggestive of a compound related to bisabolol oxides. Peaks that resulted in database match lower than 85% were considered as unknown, and corresponding fractions were subsequently purified for further identification. Compounds 2 and 5 were isolated from the combined fractions A9−A13, while fractions 15 and 18 contained compounds 1 and 4, respectively. For a detailed isolation procedure, refer to the Isolation Scheme, Supporting Information. The ethyl acetate (52.2 g) and n-butanol (39.7 g) fractions derived from liquid−liquid partitioning were combined based on their similar LC-MS profiles and purified by gravimetric CC on 850 g of silica gel (gradient elution from 95:5 CHCl3−MeOH to 5:5 CHCl3−MeOH). Ten main fractions, B1 to B10, were obtained. Fraction B5 (6.7 g) was further purified repetitively using both a Sephadex LH-20 column with MeOH 100% and RP-C18 Flash cartridges (gradient elution from MeOH−H2O 6:4 to MeOH−H2O 8:2). Compound 3 was isolated as a dark brown, amorphous solid (25.7 mg). Enzymatic Hydrolysis of 15-Hydroxybisabolol Oxide A β-DGlucoside (3). Compound 3 was hydrolyzed enzymatically as described by Xia et al.31 A 3 mg amount of compound was dissolved in ethyl acetate (0.3 mL) and added to a solution of β-glucosidase (3.2 mg of enzyme dissolved in 0.9 mL of 0.1 M DHPCA buffer, pH 5.2). The resulting solution was stirred constantly at 37 °C for 24 h and then extracted three times with ethyl acetate. The organic phase was separated, concentrated, and analyzed by GC-MS.

the presence of the carbonyl group at C-5 and not at C-6. Thus, other enzymatic steps could be involved in this case, such as allylic oxidation, epoxidation, and ring closure. Sesquiterpene biosynthesis in M. chamomilla derives from a complex “cross-talk” between the mevalonic acid metabolism and the mevalonate-independent methylerythritol phosphate metabolism.30 The enzymes involved in the cascade can be localized in the plastids, in the cytosol, or in both, thus forming a complex pathway that is not yet fully characterized. The identification of new, minor bisabolane oxidative metabolites herein reported could be useful in better understanding the biological pathways involved in sesquiterpene synthesis in German chamomile. Even though many compounds have been identified from this species, the GC-assisted fractionation approach enabled the identification of new bisabolol oxide derivatives. In addition to the identification of new compounds, this approach might serve as an easy and robust tool for rapid identification of minor compounds from dietary supplements containing German chamomile. Interestingly, many bisabolanederived compounds are known to exhibit biological properties.12−18 Bisabolol oxide A glycoside (4) has been patented for the treatment of diabetes,21 and further evaluation of 3 could be of potential interest. It is also worth noting that, because of their high polarity, bisabolol glycoside derivatives could occur in aqueous extracts of chamomile (like herbal tea preparations) and may be hydrolyzed in vivo to release the active aglycone, thus contributing to the beneficial effects of other glycoside derivatives (mainly flavonoids) occurring in aqueous preparations of M. chamomilla.

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EXPERIMENTAL SECTION

General Experimental Procedures. Specific rotations were measured on a Rudolph Research Analytical digital polarimeter at 589 nm and 20 °C, using a 5 cm path length microcell. IR spectra were recorded on Bruker Tensor 27 FT-IR and MIRacle ATRFT-IR spectrometers (Bruker Optics) or using a universal attenuated total reflection sampling accessory with a diamond ATR (attenuated total reflectance) on an Agilent Cary 630 FT-IR spectrometer. 1H (400 MHz) and 13C (100 MHz) NMR spectra were recorded on American Varian Mercury Plus 400 NMR spectrometers. 1H (500 MHz) and 13C (126 MHz) NMR spectra were recorded on a Bruker AVANCE DRX spectrometer in CDCl3 if not stated differently. Chemical shifts are referenced to the residual solvent signal (CDCl3: δH 7.26 ppm, δC 77.1 ppm). Homonuclear 1H connectivities were determined from 2DCOSY experiments. One-bond heteronuclear 1H−13C connectivities were determined using gradient-HMQC experiments in which the interpulse evolution period was optimized to 3.45 μs. Two- and threebond 1H−13C connectivities were determined by gradient-HMBC experiments in which the evolution period for long-range 1H−13C coupling constants was optimized for a 2,3JC,H of 8 Hz. Through-space 1 H connectivities were evidenced using a NOESY experiment with a mixing time of 0.450 s. Accurate masses were obtained with an HRESIMS (Agilent 6200 Series, ESI source model #G1969A equipped with TOF, Agilent Technologies). The positive-ionization mode was performed with a capillary voltage of 4000 V. Nitrogen was used as the nebulizing gas (30 psi) as well as the drying gas at 11 L/min at a temperature of 350 °C. The voltage of the PMT, fragmentor, and skimmer were set at 850, 100, and 60 V, respectively. Full-scan mass spectra were acquired from m/z 100−1000. Data acquisition and processing was done using Analyst QS software (Agilent Technologies). GC analyses were performed with an Agilent 7890A GC and an Agilent 5975C Inert XL mass selective detector, equipped with an Agilent 7693 autosampler. The injector temperature was set to 250 °C. The oven temperature program was as follows: initial temperature was 45 °C (held for 2 min), then 1.5 °C/min programmed to 100 °C, 2 °C/min programmed to 200 °C, and 10 °C/min programmed to the D

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Table 1. NMR Spectroscopic Data for the Bisabolol Oxide Derivatives 1−3 1 (CDCl3)

a

position

δC, typea

1a 1b 2 3 4a 4b 5 6a 6b 7a 7b 8a 8b 9 10a 10b 11a 11b 12 13 14 15 1′ 2′ 3′ 4′ 5′ 6′a 6′b

74.4, C 70.5, CH 24.8, CH2 23.3, CH2 75.1, C 48.2, CH 19.9, CH2 37.4, CH2

2 (CDCl3)

δH (J in Hz)b

3.46, dd (2.6, 4.3) 1.80−1.75, bme 2.04, m 1.72, bme 2.09, m 1.76, 1.48, 1.82, 1.47,

me me m me

δH (J in Hz)b

δC, typec

112.4, CH2

5.23, d (17.4) 5.04, d (10.7) 5.90, dd (17.3, 10.7)

73.0, C

144.1, CH 72.1, C 54.0, CH2

21.7, CH2 2.03, me 1.85, m 2.10−2.00 bme 3.93, dd (8.18, 5.70)

37.4, CH2 27.7, CH2 86.4, CH

131.0, CH

5.89, dt (10.3, 1.8)

70.7, C

1.20, 1.28, 1.21, 1.27,

28.3, 29.4, 28.3, 27.3,

s s s s

73.6, C 41.9, CH

2.67, d (14.1) 2.59, d (14.1)

82.5, C

5.75, dt (10.3, 2.1)

CH3 CH3 CH3 CH3

80.5, CH 28.5, CH2 21.6, CH2

2.84, d (17.1) 2.72, d (17.1)

211.5, C 54.6, CH2

67.3, C 134.9, CH

27.2, 28.0, 24.7, 29.7,

3 (acetone-d6)

δC, typea

25.1, CH2 136.8, C 120.0, CH 25.5, CH2

CH3 CH3 CH3 CH3

1.53, 1.52, 1.28, 1.27,

sf sf s s

27.9, 23.5, 21.7, 64.4, 103.5, 72.9, 75.5, 69.2, 75.0, 60.5,

CH3 CH3 CH3 CH2 CH CH CH CH CH CH2

δH (J in Hz)d

3.46 2.01, me 1.90, me 1.24, me 1.74, bme 2.08, 1.96, 2.09, 2.06,

me me me me

5.58, bs 1.96, me 1.22, 1.20, 1.01, 3.86, 4.37, 3.23, 3.41, 3.32, 3.33, 3.83, 3.64,

s s s s d (7.8) m m m m m m

100 MHz. b400 MHz. c126 MHz. d500 MHz. eSignal partially overlapping. fExchangeable. HRESIMS (positive ion) m/z 253.1785, [M − OH]+ (calcd for [C15H25O3]+, 253.1799). 15-Hydroxybisabolol oxide A β-D-glucoside (3): dark brown, amorphous solid; [α]20 D +1.9 (c 0.42, MeOH); IR (film) νmax 3366, 2924, 1697, 1451, 1376, 1360, 1248, 1164, 1076, 1021, 987 cm−1; NMR data as shown in Table 1; HRESIMS (positive ion) m/z 439.2296 [M + Na]+ (calcd for [C21H36 NaO8]+, 439.2303).

Acid Hydrolysis and Determination of Absolute Configuration of the Sugars. Compounds 3 and 4 were hydrolyzed following the method described by Wang et al.32 The glucoside (3.0 mg) was dissolved in 2 N HCl (0.8 mL) and heated to 60 °C while stirring. After 1 h, the solution was cooled and neutralized with 0.2 M NaHCO3, then extracted with ethyl acetate three times. The resulting aqueous layer gave the sugar residue after complete dryness. The residue was then dissolved in anhydrous pyridine (0.1 mL), and Lcysteine methyl ester hydrochloride 0.1 M in anhydrous pyridine (0.2 mL) was added. The mixture was heated at 65 °C for 1 h under a nitrogen atmosphere. An equal volume of Ac2O was added, and heating was continued for an additional 3 h. The solution was neutralized with 2 N HCl, then extracted with ethyl acetate, dried with anhydrous MgSO4, and concentrated. The organic phase contained the acetylated thiazolidine derivatives, which were subjected to GC-MS analysis [column Agilent HP-1 (60 m × 0.25 mm, 0.25 μm), oven programmed from 200 to 280 °C at 20 °C/min, then held for 10 min, m/z range 50−550 amu, electron energy 70 eV]. The D-configuration was confirmed for the glucose residues by comparison of the retention times and mass spectra with authentic acetylated thiazolidine derivatives produced from standard D-glucose (23.28/23.58 min) and L-glucose (23.89/24.67). 9-Hydroxybisabolol oxide A (1): yellowish oil; [α]20 D +1.8 (c 0.20, CHCl3); IR (film) νmax 3393, 2970, 2932, 2872, 1699, 1558, 1541, 1522, 1507, 1457, 1396, 1169, 1021, 986 cm−1; NMR data are shown in Table 1; HRESIMS (positive ion) m/z 272.2199 [M + NH4]+ (calcd for [C15H30NO3]+, 272.2221). Seco-bisabolol oxide B (2): colorless oil; [α]20 D +15.0 (c 0.20, CHCl3); IR (film) νmax 3472, 2973, 2931, 1699, 1456, 1370, 1148, 1119, 1066, 1025, 922 cm−1; NMR data as shown in Table 1;

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ASSOCIATED CONTENT

S Supporting Information *

The isolation procedure used, 1D and 2D NMR data, EI mass spectra for compounds 1−3, experimental data for compounds 4 and 5, and mass fragmentation scheme for bisabolol oxides A and B. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Tel: +1 (662) 915-7821. Fax: +1 (662) 915-7989. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research is supported in part by “Science Based Authentication of Dietary Supplements” funded by the Food and Drug Administration grant number 1U01FD004246 and the United States Department of Agriculture, Agricultural E

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(28) Inui, T.; Wang, Y.; Nikolic, D.; Smith, D. C.; Franzblau, S. G.; Pauli, G. F. J. Nat. Prod. 2010, 73, 563−567. (29) Huo, X.; Pan, X.; Huang, G.; She, X. Synlett 2011, 8, 1149− 1150. (30) Adam, K.-P.; Thiel, R.; Zapp, J. Arch. Biochem. Biophys. 1999, 369, 127−132. (31) Xia, Q.; Xu, D.; Huang, Z.; Liu, J.; Wang, X.; Wang, X.; Liu, S. Fitoterapia 2010, 81, 437−442. (32) Wang, W.; Li, X.-C.; Ali, Z.; Khan, I. A. Chem. Pharm. Bull. 2009, 57, 636−638.

Research Service, Specific Cooperative Agreement No. 586408-2-0009. The authors are thankful to Dr. V. Raman for authentication of the botanical material and for the German chamomile picture presented in the Table of Contents. The authors thank Dr. J. F. Parcher for proofreading the manuscript and Agilent Technologies, Inc. (Santa Clara, CA, USA) for provision of the analytical instrumentation used in this study.

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